Invasive Species Compendium

Detailed coverage of invasive species threatening livelihoods and the environment worldwide


(euryhaline hydroid)



Cordylophora (euryhaline hydroid)


  • Last modified
  • 25 September 2018
  • Datasheet Type(s)
  • Invasive Species
  • Preferred Scientific Name
  • Cordylophora
  • Preferred Common Name
  • euryhaline hydroid
  • Taxonomic Tree
  • Domain: Eukaryota
  •   Kingdom: Metazoa
  •     Phylum: Cnidaria
  •       Class: Hydrozoa
  •         Order: Anthoathecata
  • Summary of Invasiveness
  • Cordylophora is referred to as a Ponto-Caspian invasive hydroid presumably originating from the Caspian and/or Black Sea, though this has yet to be confirmed by morphological and molecular analyses. This euryha...

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Colony of Cordylophora growing on dressenid mussels collected from Lake Michigan, USA. July 2003.
TitleColony growing on dressenid mussels
CaptionColony of Cordylophora growing on dressenid mussels collected from Lake Michigan, USA. July 2003.
CopyrightNadine Folino-Rorem
Colony of Cordylophora growing on dressenid mussels collected from Lake Michigan, USA. July 2003.
Colony growing on dressenid mussels Colony of Cordylophora growing on dressenid mussels collected from Lake Michigan, USA. July 2003.Nadine Folino-Rorem


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Preferred Scientific Name

  • Cordylophora Allman, 1844

Preferred Common Name

  • euryhaline hydroid

Other Scientific Names

  • Cordylophora albicola Kirchenpauer, 1861
  • Cordylophora americana Leidy, 1870
  • Cordylophora annulata Motz-Kossowska, 1905
  • Cordylophora caspia (Pallas, 1771)
  • Cordylophora dubia Hargitt, 1924
  • Cordylophora fluviatilis Hamilton, 1928
  • Cordylophora japonica Itô, 1951
  • Cordylophora lacustris Allman, 1844
  • Cordylophora lacustris var. otagoensis Fyfe, 1929
  • Cordylophora mashikoi Itô, 1952
  • Cordylophora pusilla Motz-Kossowska, 1905
  • Cordylophora solangiae Redier, 1967
  • Cordylophora whiteleggi von Lendenfeld, 1886

International Common Names

  • English: European fouling hydroid; freshwater hydroid; Ponto-Caspian hydroid

Local Common Names

  • Austria: Keulenpolyp
  • Denmark: Brakvands-kollepolyp
  • Estonia: jarvetolvik
  • Finland: murtovesipolyyppi
  • Germany: Keulenpolyp
  • Lithuania: Kordylofora
  • Netherlands: brakwaterpoliep
  • Sweden: Klubbpolyp

Summary of Invasiveness

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Cordylophora is referred to as a Ponto-Caspian invasive hydroid presumably originating from the Caspian and/or Black Sea, though this has yet to be confirmed by morphological and molecular analyses. This euryhaline hydroid occurs in fresh and brackish habitats globally with an expanding distribution due to increased ship transport (via hull and ballast water). The spread and establishment seems to be enhanced by the hydroid's physiological ability to acclimate and proliferate in a wide range of salinities (Folino, 2000; bij de Vaate et al., 2002; Janssen et al., 2005). Successful establishment is strengthened via a dormant stage (menont) (Roos, 1979) that has notable regeneration capabilities in varying salinities (Kinne, 1958; N Folino-Rorem, Wheaton College, Illinois, USA, personal communication, 2009). Definitive records for this hydroid as an invasive species are vague though Roch (1924) presents a thorough global summary of documented locations. Cordylophora is not on the IUCN alert list.

Taxonomic Tree

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  • Domain: Eukaryota
  •     Kingdom: Metazoa
  •         Phylum: Cnidaria
  •             Class: Hydrozoa
  •                 Order: Anthoathecata
  •                     Family: Oceanidae
  •                         Genus: Cordylophora

Notes on Taxonomy and Nomenclature

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The taxonomy of this hydroid at the family and genus level is in a tentative state. Some (Cartwright et al., 2008; Folino-Rorem et al., 2009a) use the provisional family name of Oceaniidae Eschscholtz, 1829 proposed by Schuchert (2004, 2005) whereas others (Calder and Kirkendale, 2005; Jankowski et al., 2008) use the family name Cordylophoridae von Lendenfeld, 1885. Furthermore, other researchers (Bouillon et al., 2006; Stepanjants et al., 2006) use Clavidae McCrady, 1859. Using Oceaniidae or the more historically accurate name of Cordylophoridae at this point in time are recommended until more morphological and molecular data are available to clarify the family status (D Calder, Centre for Biodiversity & Conservation Biology, Royal Ontario Museum, Canada, personal communication, 2009; P Schuchert, Museum d'Histoire Naturelle, Geneva, Switzerland, personal communication, 2009).

A consensus for species in the genus Cordylophora is difficult and lacking due in part to the high degree of morphological plasticity and taxonomic history. The historical and therefore perhaps erroneous identification of specimens (e.g., ascribing a new or different name when found in a new location (Carlton, 2009)) that appear to be Cordylophora has led to confusion in this hydroid’s taxonomy. The following species have been published (in order by year proposed) although it is somewhat unclear how many are definitive species: C. caspia (Pallas, 1771), C. lacustris (Allman, 1844), C. albicola (Kirchenpauer, 1861), C. americana (Leidy, 1870), C. whiteleggi (von Lendenfeld, 1886), C. annulata (Motz-Kossowska, 1905), C. pusilla (Motz-Kossowska, 1905), C. dubia (Hargitt, 1924), C. fluviatilis (Hamilton, 1928), C. lacustris var. otagoensis (Fyfe, 1929), C. japonica (Itô, 1951), C. mashikoi (Itô, 1952), C. sloangiae (Redier, 1967), and C. inkermanica (Marfenin, 1983)(see Hamilton, 1883; Fyfe, 1929; Itô, 1951, 1952; Calder, 1988; Folino, 2000; Schuchert, 2004).  More recently, several of these species are considered synonyms with Cordylophora caspia (C. albicola, C. americana, C. lacustris,C. lacustris var. otagoensis, and C. whiteleggi) though no type material exists for C. caspia (Schuchert, 2004).

This invasive hydroid was initially named Tubularia (spelled Tvbvlaria) caspia then combined with the genus Cordylophora as C. caspia (Pallas, 1771). In 1844, Allman described C. lacustris, a species later proposed as a synonym to C.caspia (Roch, 1924). In addition, Cordylophora fluviatilis (Hamilton, 1928) is considered to be synonymous with C. caspia since Fyfe (1929) and Briggs (1931) equate C. fluviatilis with C. lacustris. Most authors have adopted Roch’s (1924) proposed synonymy for Cordylophoracaspia and Cordylophoralacustris, whereas others believe that although they appear morphologically identical, they are in fact different species with different habitat preferences, C. caspia being a brackish water species and C. lacustris being freshwater (Folino, 2000; Schuchert, 2004). In addition, Hargitt (1924) suggested that C. dubia was also synonymous with C. lacustris. Other previously proposed species of Cordylophora are equated with other genera (C. annulataMotz-Kossowska, 1905 =Pachycordyle napolitana; C. inkermanica Marfenin, 1983 = Pachycordyle navis; C. pusilla = Pachycordyle pusillaMotz-Kossowska, 1905) (Calder, 1988; Stepanjants et al., 2000; Schuchert, 2004, respectively). If the previous taxonomic reclassifications are accepted, there are four remaining, recognized species in the literature for the genus Cordylophora: C. caspia, C. japonica, C. mashikoi and C. solangiae. These remaining species have yet to be assessed closely to confirm their distinctiveness or whether they are synonyms.

Recent molecular results support the existence of multiple species of Cordylophora based on their occurrence in freshwater versus brackish habitats (Folino-Rorem et al., 2009a). How these potentially new species are assimilated into the current taxonomic system has yet to be determined. No doubt, the taxonomy of the genus Cordylophora is in need of a thorough and extensive morphological and molecular analysis in light of eco-plasticity.

Potentially related genera/species are presented though, again taxonomic uncertainty is present.  The genus Cordylophora resembles the genus Pachycordyle and the placement of these two genera in the same or different families or even stating that they are synonyms (Morri, 1980, 1981) is under dispute(see review by Schuchert (2004)). Vervoort (1964) found Bimera baltica Stechow, 1927 indistinguishable from C. caspia. In addition, Velkovrhia enigmatica is very similar to C. caspia and may simply be an ecomorph of C. caspia (Schuchert, 2007).



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Cordylophora is a colonial, athecate euryhaline hydroid occurring in freshwater and brackish habitats globally (Roos, 1979; Folino, 2000). This is one of the few known freshwater Cnidaria and appears be the only freshwater, colonial hydrozoan (Clarke, 1878). Colonies grow on living and non-living hard substrata, with colonies ranging in height from 1-12 cm, though this varies depending on habitat/salinity (Kinne, 1958; Arndt, 1984). Although colony growth is optimal at 15-17 psu at 20°C, this hydroid can tolerate a wide range of salinities (Roch, 1924; Kinne, 1956; Arndt, 1984). Colonies consist of polyps (gastrozooids or hydranths) specialized for feeding or reproduction (gonophores that produce sporosacs) (Allman, 1844; Fraser, 1944). The tentacles on the hydranths use microbasic euryteles and desmonemes nematocysts for capturing prey (Itô, 1951; Stepanjants et al., 2000; Schuchert, 2004). An upright or hydrocaulus can be unbranched or monopodially branched with a terminal hydranth or feeding polyp. Hydranths bear scattered filiform tentacles with a conical hypostome; the number of tentacles typically ranges between 14 and 16 though some can have as many as 27 but this varies relative to salinity (Kinne, 1958; Schuchert, 2004). Colonies sometimes have annulations of the perisarc at the base of an upright or side branch (Calder, 1968; Gosner, 1971; P Schuchert, Museum d'Histoire Naturelle, Geneva, Switzerland, personal communication, 2004). Historically, it is the number, type and arrangement of tentacles on the hydranth that are the primary diagnostic features used in identifying this genus.

Colonies are dioecious possessing either male or female sporosacs (or gonophores) for sexual reproduction. Fertilized eggs develop into free-swimming planula larvae that settle to form a sessile, primary polyp. A medusa stage is lacking (Allman, 1853). Colonies reproduce asexually by budding. Cordylophora survives cold temperatures and periods of unfavourable conditions via small masses of tissues, called menonts, which remain in the perisarc of the hydrocauli and/or uprights. This tissue regenerates when temperatures increase or conditions become more favorable (Kinne, 1971; Roos, 1979; Folino, 2000).


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Cordylophora occurs on all continents except Antarctica (see pictures). This hydroid is absent from fully marine habitats though colonies demonstrate suboptimal growth in laboratory cultures at higher salinities (30-40 psu) (Kinne, 1958).

In addition to the countries listed in the Distribution Table, Cordylophora is also present in Iceland (Lake Ljosavatn) (R Campbell, University of California, USA, personal communication, 2009) and Thailand (Lam Takong Reservoir, Khorat) (T Wood, Wright State University, Ohio, USA, personal communication, 2009). As well as the records in the table, there are additional reports from: Germany (Ryck River, Greifswald), Ireland (Shannon River, Limerick) and the Netherlands (River Waal, Nijemegen) (N Folino-Rorem, Wheaton College, Illinois, USA, personal communication, 2009);  the USA: California (Napa and Petaluma rivers, San Francisco Bay Area), Illinois (Illinois and Des Plaines rivers, LaSalle Lake, several harbours in Lake Michigan), Massachusetts (Merrimack River, Amesbury), Michigan (Lake Michigan, Muskegon), New Hampshire (Lamprey River, Newmarket; Squamscott River, Exeter; Jackson Landing, Durham), New York (Finger Lakes, Cayuga and Seneca Lakes, Lake Ontario, Rochester), Pennsylvania (Lake Erie, Presque Isle, State Park), Virginia (James River, Jamestown Settlement) (N Folino-Rorem, Wheaton College, Illinois, USA, personal communication, 2009) and West Virginia (Floyd Lake, Harrison County) (R Campbell, University of California, USA, personal communication, 2009); and the United Kingdom (Wraysbury, Middlesex) (R Mant, University of Cambridge, UK and N Folino-Rorem, Wheaton College, Illinois, USA, personal communication, 2009).


Distribution Table

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The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.

Continent/Country/RegionDistributionLast ReportedOriginFirst ReportedInvasiveReferenceNotes


AzerbaijanPresent1969Native Not invasive Naumov, 1969; Stepanyants, 2009North, Middle and South Caspian Sea
ChinaPresentPresent based on regional distribution.
-AnhuiPresent1951Introduced Invasive Roch, 1924Tai Hu
-ShanghaiPresent1981Introduced Invasive Huang et al., 1981Brackish-freshwater: Changjiang River Estuary
IraqPresent1984Introduced Invasive Arndt, 1984Brackish, Shatt Al-Arab
JapanPresentPresent based on regional distribution.
-ShikokuPresent1951Introduced Invasive Itô, 1951Brackish creek in Kôchi
PhilippinesPresent1924Introduced Invasive Hargitt, 1924Mololas River


EgyptPresent2009Introduced Invasive Roch, 1924; Dumont, 2009; Green and El-Moghraby, 2009Brackish, Lake Qarun in Fayum
SudanPresent1949Introduced Invasive Rz?ska, 1949White Nile River, Tongo, Malakal, Bahr El Ieraf

North America

CanadaPresentPresent based on regional distribution.
-British ColumbiaPresent1979Introduced Invasive Mace and Mackie, 1970; Carlton, 1979Brackish, Albert Head Lagoon
-QuebecPresent1971Introduced Invasive Calder, 1971
USAPresentPresent based on regional distribution.
-ArizonaPresent1989Introduced Invasive Pennak, 1989Freshwater
-ArkansasPresent1989Introduced Invasive Roch, 1924; Pennak, 1989Freshwater, Arkansas River, Little Rock
-CaliforniaPresent2006Introduced Invasive Hand and Gwilliam, 1951; Hand and Jones, 1957; Aldrich, 1961; Siegfried et al., 1980; Cohen and Carlton, 1995; Wasson et al., 2005Lake Merced
-ConnecticutPresent2002Introduced Invasive Smith et al., 2002Connecticut River, Essex
-FloridaPresent1996Introduced Invasive Streever, 1992; Havens et al., 1996Freshwater, Little River Spring cave system
-HawaiiPresentIntroduced Invasive Cooke, 1977Brackish, Cape Kinau, Maui
-IllinoisPresent2009Introduced Invasive Smith, 1910; Lipsey and Chimney, 1978; Pennak, 1989Freshwater Quiver Lake Havana
-KansasPresentIntroduced Invasive Ransom, 1981Freshwater, Melvern Lake, Osage County
-KentuckyPresent1961Introduced Invasive Garman, 1922; Hargitt, 1923; Weise, 1961Freshwater
-LouisianaPresentIntroduced Invasive Roch, 1924; Porrier and Denoux, 1973; Koetsier and Bryan, 1989Freshwater, Mississippi River, W. Feliciana Parish Louisiana
-MarylandPresent2005Introduced Invasive Clarke, 1878; Bibbins, 1892; Hargitt, 1897; Cory, 1967; Cory and Nauman, 1969; Jewett, 2005Curtis Creek, Baltimore
-MassachusettsPresent2002Introduced Invasive Hyatt, 1866; Nutting, 1901; Hargitt, 1908; Blake, 1932; Fulton, 1962; Smith et al., 2002Brackish water, Mystic Pond
-MinnesotaPresent2003Introduced Invasive Grigorovich et al., 2003Lake Superior, Duluth
-New HampshirePresent2008Introduced Invasive Gaulin et al., 1986; Harris and Dijkstra, 2007Great Bay Estuary
-New JerseyPresent1989Introduced Invasive Dean and Haskin, 1964; Pennak, 1989Raritan River
-New YorkPresent2008Introduced Invasive Mills et al., 1996; Walton, 1996; Strayer and Malcom, 2007Oligohaline, Hudson River
-North CarolinaPresent1975Introduced Invasive Dean and Bellis, 1975Pamlico River Estuary
-OhioPresent1964Introduced Invasive Davis, 1957; Hubschman and Kishler, 1972Chagrin Harbour, Lake County
-OklahomaPresent1968Blair, 1964; Ransom, 1968Filtration plant, Mohawk Park, Tulsa
-OregonPresent1967Introduced Invasive Haertel and Osterberg, 1967Columbia River Estuary; Harrington Pt. Astoria & Chinook Pt.
-PennsylvaniaPresent2007Introduced Invasive Leidy, 1870; Potts, 1884Schuylkill River, Fairmont
-Rhode IslandPresent1951Introduced Invasive Leidy, 1870Newport, RI
-South CarolinaPresent1976Introduced Invasive Calder, 1976Estuarine areas of South Carolina
-TennesseePresent1989Introduced Invasive Isom and Sinclair, 1962; Delong and Payne, 1985; Smith and Hamilton, 2004Tennessee River, Cumberland River & Duck River
-TexasPresentIntroduced Invasive McClung et al., 1978; Davis, 1980Pecos River at Orla, Girvin, Sheffield and Shumla
-VirginiaPresent2007Introduced Invasive Schmitt, 1927; Calder, 1968; Calder, 1971Great Falls, Potomac River
-WashingtonPresent1998Introduced Invasive Cohen et al., 1998; Mills, 1998Brackish, Edison, near mouth of Samish River

Central America and Caribbean

PanamaPresent2001Introduced Invasive Hildenbrand, 1939; Jones and Rutzler, 1975Gatún Locks, Panama Canal

South America

ArgentinaPresent1957Introduced Invasive Davis, 1957Rio de la Plata, Argentina
BrazilPresentPresent based on regional distribution.
-Mato Grosso do SulPresent2004Introduced Invasive Roque et al., 2004Paraná River
-Minas GeraisPresent2004Introduced Invasive Roque et al., 2004Paraná River
-Rio de JaneiroPresent1951Introduced Invasive Roch, 1924Material from Van Beneden Zoological Institute of Berlin
-Sao PauloPresent2004Introduced Invasive Roque et al., 2004Paraná River
ChilePresent2007Introduced Invasive Galea, 2007Fjords region of southern Chile; fjord Comau


AustriaLocalisedIntroducedNOBANIS, 2011Cordylophora caspia
BelgiumPresent1972Introduced Invasive Roch, 1924; Rose and Burnett, 1969; Lesh-Laurie, 1972Canal near Ostend
DenmarkPresent1924Introduced Invasive Roch, 1924
EstoniaPresent2004Introduced Invasive Roch, 1924; Ojaveer et al., 2004
FinlandPresent1994Introduced Invasive Roch, 1924; Arndt, 1984; Jormalainen et al., 1994Helsinki, Raseborg and Finnish Inlets
FrancePresent2005Introduced Invasive Roch, 1924; Devin et al., 2005Aquarium at the Garden of Plants
GermanyPresent2006Introduced Invasive Roch, 1924; Kinne, 1956; Tessnow, 1959; Calmano et al., 1992; Ringelband and Karbe, 1996; Nehring, 2006Several locations: Elbe River, Bille River, Travellodge Bay, near Berlin
HungaryPresent2008Introduced Invasive Muskó et al., 2008Freshwater, Lake Balaton and Danube River
IrelandPresent2004Introduced Invasive Allman, 1844; Roch, 1924Docks of the Grand Canal, Dublin
ItalyPresent1979Introduced Invasive Morri, 1979Lake of Fondi
LithuaniaPresent2005Native Not invasive Roch, 1924; Olenin and Daunys, 2005Coast along Baltic Sea
LuxembourgPresent1987Introduced Invasive Massard and Geimer, 1987The Grand - Duchy
NetherlandsPresent2007Introduced Invasive Roch, 1924; Vervoort, 1964; Wolff, 1999Several cities and canals
NorwayPresent2002Introduced Invasive Hopkins, 2002; Leppäkoski et al., 2002
PolandPresent1984Introduced Invasive Arndt, 1984Martwa Wisla
Russian FederationPresentPresent based on regional distribution.
-Southern RussiaPresent2006Native Not invasive Roch, 1924; Shokin et al., 2006Black, Caspian and Azov Seas
SpainPresent2007Introduced Invasive Roca, 1987; Escot et al., 2007Balearic Water
SwedenPresent1996Introduced Invasive Clarke, 1878; Roch, 1924; Franzen, 1996Stockholm and surrounding areas
UKPresent1987Introduced Invasive Roch, 1924; Moore, 1952; Chain, 1980; Barnes, 1987London , Norfolk, Tipton Canal
UkrainePresent2007Native Not invasive Alexandrov et al., 2007Black and Azov Seas


AustraliaPresentPresent based on regional distribution.
-New South WalesPresentIntroduced Invasive Roch, 1924Parramata River, near Sidney
-TasmaniaPresentIntroduced Invasive Roch, 1924The Nebenflus River between Wyngard and Flowerdale
-Western AustraliaPresent2005Introduced Invasive Halse et al., 2002; Pinder et al., 2005Coyrecup Wetland
New ZealandPresent1998Introduced Invasive Roch, 1924; Cranfield et al., 1998Esk River, Hawke’s Bay

History of Introduction and Spread

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The global spread and establishment of C.caspia (a Ponto-Caspian invasive species from the Black, Caspian, Azov Seas and surrounding areas) is primarily attributed to increased ship transport through canals and rivers via ship ballast and/or hull fouling (Folino, 2000; bij de Vaate et al., 2002; Pienimäki and Leppäkoski, 2004; Janssen et al., 2005; Streftaris et al., 2005). The hydroid presumably originated in the Black and/or Caspian Seas, spreading west via a northern corridor to the Baltic Sea, a north central corridor toward the North Sea and a southern corridor toward the Mediterranean Sea (bij de Vaate et al., 2002; Ketelaars, 2004). The avenues of rivers and canals to Europe facilitated the establishment in the coastal/estuary areas especially along the North and Baltic Seas. A summary of the spread of C. caspia suggests its presence in the North Sea in 1858 with records in the Rhine River in 1874 (van Riel et al., 2006). It is unclear if the hydroid was present in the Baltic Sea before the North Sea or vice versa. Streftaris et al. (2005) document the occurrence of C. caspia in the Baltic at 1883 and the North Sea in 1884. Additional records document this hydroid in the Baltic Sea in 1870 and being present in 1899 in the Kiel Canal (the canal connecting the North and Baltic Sea) (Gollasch and Rosenthal, 2006). Nevertheless, once established, this hydroid spread to European countries with records of C.caspia in French freshwater systems around 1970 (Devin et al., 2005)

Roch (1924) records the first accounts for Cordylophora in Asia, Africa, and Australia. The earliest record of this hydroid in Central America is from the Panama Canal (Hildenbrand, 1939), obviously transported by ships as a fouler or in ballast water (Cohen, 2006). The first records in North America are for Massachusetts in 1860 (Verrill et al., 1873), while it was later discovered on the west coast of Puget Sound and the San Francisco Bay areas circa 1920 (Cohen et al., 1998; Ruiz et al., 2000; Wonham and Carlton, 2005). Records for Cordylophora (lacustris) in the midwest United States (specifically Kentucky) were published by Garman (1922). Davis (1957) found Cordylophora (lacustris) in the Chargin River in Ohio while Cordylophora (lacustris) was well established in Western Lake Erie by 1972 (Hubschman and Kishler, 1972). Cordylophora spp. supposedly made its way into the Great Lakes via the St Lawrence River System in 1956 (Mills et al., 1993).

Risk of Introduction

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The risk of Cordylophora being introduced and/or spreading seems very likely based on its worldwide distribution and changes in water quality due to anthropogenic activity that alters salt concentrations (Hubschman, 1971; Folino, 2000). In addition, the likelihood of new introductions continues with increased shipping (spread via ship ballast and/or hull fouling) (Folino, 2000; bij de Vaate et al., 2002; Pienimäki and Leppäkoski, 2004; Janssen et al., 2005; Streftaris et al., 2005).


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Because this hydroid is a euryhaline organism, it occurs in both freshwater and brackish habitats (0-32 psu). Whether there are species differences relative to salinity tolerances is yet to be determined (Folino-Rorem et al., 2009a). Colonies are present in tidal areas, rivers, lakes, lagoons and ponds and grow on bivalve shells (e.g., Dreissena, Anodon)(Allman, 1853; Schulze, 1871; Folino-Rorem and Stoeckel, 2006), submerged vegetation (Fyfe, 1929; Roos, 1979; Strayer and Malcom, 2007; El-Shabraway and Fishar, 2009), rocks, wooden pilings, living crab and gastropod shells and floats (Fraser, 1944; Roos, 1979; Gaulin et al., 1986; Barnes, 1994; bij de Vaate et al., 2002; Kautsky, 2008). Estuarine colonies of Cordylophora also occur on eelgrass blades (Chester, 1996), and grow on barnacles (N Folino-Rorem, Wheaton College, Illinois, USA, personal communication, 2009) and oyster Crassostrea virginica shells (Calder, 1971; Carlton, 1979).

Habitat List

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Inland saline areas Principal habitat Harmful (pest or invasive)
Inland saline areas Principal habitat Productive/non-natural
Estuaries Principal habitat Harmful (pest or invasive)
Estuaries Principal habitat Productive/non-natural
Lagoons Principal habitat Harmful (pest or invasive)
Lagoons Principal habitat Productive/non-natural
Coastal areas Present, no further details Productive/non-natural
Lakes Principal habitat Harmful (pest or invasive)
Lakes Principal habitat Productive/non-natural
Reservoirs Principal habitat Harmful (pest or invasive)
Reservoirs Principal habitat Productive/non-natural
Rivers / streams Principal habitat Productive/non-natural
Ponds Principal habitat Productive/non-natural

Biology and Ecology

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The karyotype for Cordylophora is n=30 (Stepanjants et al., 2000). The following nucleotide sequences for several populations of Cordylophora spp. are available at GenBank at partial sequence; Cartwright et al.: 16S ribosomal RNA gene, partial sequence; mitochondrial; Evans et al. Sch485 28S large subunit ribosomal RNA gene, partial sequence; Folino-Rorem et al: 28S large subunit ribosomal RNA gene. Sequences for Cordylophora mitochondrial 16S rDNA are available at the Hydrozoan Taxonomy site (PEET: Partnerships for Enhancing Expertise in Taxonomy) at:

The molecular analyses of several Cordylophora populations collected worldwide are addressed using DNA sequence data from two mitochondrial loci [the small subunit 16S rRNA and cytochrome c oxidase subunit I (COI)] and one nuclear locus (28S large nuclear rRNA). Results suggest multiple cryptic species within the genus (Folino-Rorem et al., 2009a). Additional sequences for 18S and 28S are presented in an assessment of the phylogenetic placement of Polypodium hydriforme (Evans et al., 2008).

Reproductive Biology

Colonies are dioecious possessing either male or female gonophores arising from either hydranth pedicels or from branches below a given hydranth (Allman, 1853; Fyfe, 1929). One source, Vervoort (1964) states that male and female gonophores occur on the same colony in C. caspia. In sexual reproduction a male sporosac (white) releases sperm into the water with the eggs being fertilized within female sporosacs (pinkish/purplish). The fertilized eggs develop into free-swimming planula larvae; a medusa stage is lacking. The female sporosacs release free-swimming planulae that eventually settling to form a sessile, primary polyp. The primary polyp then buds asexually forming a colony (See Pictures).

Colonies reproduce asexually by budding with prolific growth during the summer and early autumn months (Roos, 1979; Jormalainen et al., 1994; Muskó et al., 2008; N Folino-Rorem, Wheaton College, Illinois, USA, personal communication, 2009). Brackish colonies of Cordylophora (caspia) demonstrated asexual reproduction right after winter dormancy (menont stage) in the beginning stages of colony development (Cory, 1967; Roos, 1979; Jormalainen et al., 1994). Profuse colonies are found in both freshwater and brackish habitats from June-September or October (depending on temperature, food availability and presence or absence of predators) (Jormalainen et al., 1994). The presence of sexually mature colonies is greatest in June and July (brackish) (Cory, 1967; Jormalainen et al., 1994); freshwater (Fyfe, 1929; Muskó et al., 2008; N Folino-Rorem, Wheaton College, Illinois, USA, personal communication, 2009).

Physiology and Phenology

Similar to other invasive Ponto-Caspian organisms, Cordylophora often contends with changes in salinity when colonizing a new habitat (Ricciardi and Rasmussen, 1998; bij de Vaate et al., 2002; Paavola et al., 2005). The published native salinity range for a brackish population of Cordylophora (caspia) from West Germany is 0-35 psu at 20°C, with 15-17 psu being the optimal range (Roch, 1924; Kinne, 1958; Arndt, 1984). Chester et al. (2000) observed optimal growth for a brackish population of Cordylophora (lacustris) at 20 psu, 25°C. Similarly, a brackish population of Cordylophora (lacustris) was cultured for 20 days at 6, 12, 18 and 24 psu at 20°C; the colonies at 6 psu demonstrated the greatest growth rate (Blezard, 1999). It was found that a freshwater population transitioned to 24 psu demonstrated the highest growth rate at 4 psu while a brackish population transitioned from 10 psu to 24 psu and from 10 psu to 0 psu demonstrated higher growth rates between 12 and 18 psu at 22°C (N Folino-Rorem, Wheaton College, Illinois, USA, personal communication, 2009). In general, it is difficult to propose an optimal salinity since various temperatures have been used in laboratory studies and endogenic differences exist for an organism with ecologically plastic physiological responses to salinity and temperature (Arndt, 1984).

Colony growth form in conjunction with hydranth size, tentacle length and number, and hydranth cell numbers, nematocyst size and number and size of nuclei vary or are plastic with variations in salinity and temperature (Kinne, 1958; 1971). When a brackish population is gradually transferred to either freshwater (0 psu) or sea water (30 psu), the hydranths become shorter in length and wider in width while the tentacles are shorter. Brackish (15 psu) colonies have hydranths that are longer in length and narrower in width, with longer tentacles compared to colonies in fresh and salt water (Kinne, 1958). Overall form varies with colonies reared in fresh and saltwater; colonies are also shorter and less branching, while colonies reared in brackish water are taller and more branching (Kinne, 1958). At the tissue level, the following demonstrate a decreased range in salinity tolerance respectively: stolon material, hydranth tissue, tentacles and lastly gonosphores (Kinne, 1958). To support these findings, menont tissue present in the stolons of a freshwater population are able to regenerate when grown in 0, 2, 4, 8 and 12 psu (N Folino-Rorem, Wheaton College, Illinois, USA, personal communication, 2009). Furthermore, the size and number of gonophores vary with salinity: a brackish population demonstrates maximum gonophore production between 5-15 psu. In addition, the number of eggs/gonophore is reduced at suboptimal and superoptimal salinities (Kinne, 1971).

Cordylophora digestion times vary with salinity. Brackish population feeding times are shortest at 16.7 psu while those in freshwater or 30 psu are greater. This same pattern is true for ingestion time (prey capture and prey within the gut) (Kinne and Paffenhöfer, 1965). The ability to capture prey may vary relative to salinity since the length and width of nematocysts are greater in freshwater and smaller in saltwater (Kinne, 1958; 1971). This is in contract to Kinne's (1958) findings with regards to hydranth size relative to salinity where hydranths are larger in brackish water. This suggests that salinity may affect the functionality of nematocysts and therefore perhaps feeding abilities in different habitats.

Cordylophora respiration rates (oxygen consumption) are affected by temperature and salinity (Arndt, 1984). Respiration rates for a brackish population of Cordylophora (caspia) are lower at 10°C compared to 20°C at three salinities of 1, 10 and 24 psu while respiration was higher at 1 and 24 psu (Arndt, 1984).
As previously noted, the extensive phenotypic and physiological plasticity of Cordylophora is a major difficulty in trying to clarify the taxonomic status of this organism. The fundamental question is whether several species exist relative to these morphological and physiological differences or is there extensive genetic variation with Cordylophora being a truly euryhaline taxon capable of flourishing in a wide range of salinities. Alternatively, there may exist multiple species of Cordylophora with different habitat preferences (Folino-Rorem et al., 2009a).

Most studies present aspects of the seasonality of Cordylophora but few address complete years of life stage information (Allman, 1853; Cory, 1967; Jormalainen et al., 1994; Muskó et al., 2008). As temperatures decrease during late autumn, fewer unprights with hydranths are present and colonies enter the dormancy or menont stage (Roos, 1979; Muskó et al., 2008; N Folino-Rorem, Wheaton College, Illinois, USA, personal communication, 2009). In both freshwater and brackish populations in temperate locations, colonies begin to come out of dormancy and regenerate hydranths in the spring during April-May relative to temperature (between 6 and 15°C). Colony growth increases with temperature and gonophores are produced when temperatures reach 10°C in brackish (Arndt, 1989) or around 15-18°C in freshwater (Muskó et al., 2008; N Folino-Rorem, Wheaton College, Illinois, USA, personal communication, 2009). Depending on the population, extreme salinities can alter or suppress gonophore production (Kinne, 1971). Gonophores (and therefore sporosacs) for sexual reproduction are present in June and are most abundant from July-October (Allman, 1853; Cory, 1967; Jormalainen et al., 1994; Muskó et al., 2008; N Folino-Rorem, Wheaton College, Illinois, USA, personal communication, 2009).


Cordylophora is a carnivorous suspension raptorial feeder andpreys primarily on zooplankton such as copepods, cladocerans (Roos, 1979; Olenin and Leppäkoski, 1999; Folino-Rorem and Stoeckel, 2006) and ostracods (Hargitt, 1897). Cordylophora (caspia) has even been observed feeing on the invasive cladoceran, Cercopagis in Curonian lagoon and Lake Michigan (Gasiunaite and Didziulis, 2000; N Folino-Rorem, Wheaton College, Illinois, USA, personal communication, 2009, respectively). In addition, dreissenid mussel larvae and benthic macroinvertebrates such as worms and insect larvae (chironomids) are also consumed (Witkor, 1969; Roos, 1979; Menzie, 1981; Olenin and Leppäkoski, 1999; bij de Vaate et al., 2002; Rahel, 2002; Smith et al., 2002; Folino-Rorem and Stoeckel, 2006). Similar macroinvertebrates serve as fish prey items (Smith, 1918; Fullerton et al., 1998; Pothoven et al., 2000) and this overlap suggests a possible significant predatory role of this hydroid as a competitor for macroinvertebrate fish prey. Preliminary results suggest that Cordylophora exhibits differences in prey consumption as a function of seasonality and habitat locations such as rivers versus lakes and ponds, benthic versus the undersides of docks and freshwater versus brackish (Folino-Rorem and Stoeckel, 2006). Since nematocyst size and number vary with salinity, the ability to capture prey may vary in habitats of different salinities. Furthermore, Cordylophora hydranths are capable of capturing and ingesting large prey such as chironomid larvae that are 2-3 times larger than the hydranths (Roos, 1979; Smith et al., 2002; Folino-Rorem and Stoeckel, 2006). This unique feeding capability is accomplished by folding the larvae in half (See Pictures) or with several hydranths attacking the same chironomid demonstrating that hydranths of a given colony act to jointly capture and ingest prey, especially as prey size increases (Josephson, 1961; N Folino-Rorem, Wheaton College, Illinois, USA, personal communication, 2009). Prey behaviour (which varies with chironomid instars) and morphology seems to play a role in the ability of Cordylophora to capture and ingest prey. Chironomid thrashing may facilitate capture by Cordylophora while large posterior spines of Daphnia magna prevent Cordylophora from fully ingesting it (Folino-Rorem et al., 2009b).


Freshwater populations of Cordylophora are often associated with bryozoa (e.g. Plumatella), sponges and protozoa that colonize the stalks or perisarc of the colonies (Potts, 1884; Smith, 1910; Davis, 1957; Weise, 1961; Roos, 1979; Massard and Geimer, 1987). As might be expected, Cordylophora is sometimes found in similar habitats in conjunction with other Cnidaria such as Craspedocustasowerbyi (Garman, 1922; Hargitt, 1923; N Folino-Rorem, Wheaton College, Illinois, USA, personal communication, 2009) and Hydra (Allman, 1844; Koetsier and Bryan, 1989).

The filamentous structure of Cordylophora provides a substrate to inhabit for macroinvertebrates such as chironomids and caddis flies (Roque et al., 2004; Folino-Rorem and Stoeckel, 2006).

In addition, Cordylophora often is often found in association with dreissenid mussels by growing on theshells (Schulze, 1871; Rahel, 2002; Folino-Rorem and Stoeckel, 2006). A possible explanation for greater abundances of this hydroid is an increase in hard substrata provided by the extensive distribution of dreissenid mussels colonizing soft substrata. Dreissenid mussels provide hard substrata by colonizing soft substrata in Lake Michigan (Vanderploeg et al., 2002; Janssen et al., 2005; N Folino-Rorem, Wheaton College, Illinois, USA, personal communication, 2009). Thus, dreissenid mussels may enhance the expansion for the Ponto-Caspian associate, Cordylophora in habitats where they coexist. It is likely that they provide more hard substrata in the Great Lakes for the expanding establishment of Cordylophora. Of interest is whether Cordylophora will inhabit new habitats with the decline in Dreissena polymorpha due to replacement by Dreissena bugensis (Zhulidov et al., 2006).

Environmental Requirements

In freshwater systems, C. caspia is becoming a prevalent biofouler due to water quality changes (Folino, 2000; Smith et al., 2002; Folino-Rorem and Indelicato, 2005). Increased salts (chlorides) from runoff containing road salt seem to enhance the distribution of this hydroid (Hubschman, 1971; Smith, 1989). Additionally, salt concentrations can increase at power plant cooling lakes due to water loss from evaporative cooling also enhancing the occurrence of Cordylophora in freshwater systems (Folino-Rorem and Indelicato, 2005). Sodium and potassiumare known required elements for Cordylophora growth (Fulton, 1962). Extensive research by Roch (1924), Fulton (1962) and Kinne (1971) address the water chemistry requirements for Cordylophora growth.


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C - Temperate/Mesothermal climate Preferred Average temp. of coldest month > 0°C and < 18°C, mean warmest month > 10°C

Water Tolerances

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ParameterMinimum ValueMaximum ValueTypical ValueStatusLife StageNotes
Depth (m b.s.l.) Optimum Surface attached to docks to 15 m
Dissolved oxygen (mg/l) Optimum >2 preferred (Fulton, 1962)
Hardness (mg/l of Calcium Carbonate) Optimum Calcium is required for growth (Fulton, 1962)
Salinity (part per thousand) Optimum 15-17 PSU preferred, 0-35 PSU tolerated (Kinne, 1958)
Water pH (pH) 6.3 8.6 Optimum 5.10-9.45 tolerated (Fulton, 1962)
Water temperature (ºC temperature) Optimum 2-24 tolerated. Range influenced by salinity (Kinne, 1971)

Natural enemies

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Natural enemyTypeLife stagesSpecificityReferencesBiological control inBiological control on
Gammarus Predator Adult to genus Roos, 1979
Tenellia Predator Adult not specific Blezard, 1999; Chester, 1996; Gaulin et al., 1986; Harris et al., 1980; Jormalainen et al., 1994
Tridentiger bifasciatus Predator Matern and Brown, 2005

Notes on Natural Enemies

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A well-known predator on brackish Cordylophora coloniesis the estuarine nudibranch, Tenellia adspersa. Tenellia is a generalist nudibranch but is often feeds on brackish colonies of Cordylophora causing a noticeable decline in colonies during late summer and early autumn in temperate areas (Harris et al., 1980; Gaulin et al., 1986; Arndt, 1989; Chester, 1996; Blezard, 1999). Furhermore, predation by Tenellia adspersa (a synonym of Embletonia pallida) may graze so heavily on Cordylophora (caspia) colonies causing predator-induced dormancy (Jormalainen et al., 1994; Jewett, 2005). Blezard (1999) demonstrated that fecundity and development of Tenellia were less than optimal at salinities below 12 psu creating a salinity refuge from predation for Cordylophora (lacustris). Another invertebrate predator of Cordylophora hydranths of is the amphipod, Gamarus (Roos, 1979).

The author of this datasheet has knowledge of only one vertebrate predator on Cordylophora caspia, the invasive shimofuri goby (Tridentiger bifasciatus) in San Francisco Estuary, California (Matern and Brown, 2005).

Means of Movement and Dispersal

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The majority of citations refer to ship hulls and or ballast water as the primary means of dispersal for Cordylophora (Folino, 2000; bij de Vaate et al., 2002; Pienimäki and Leppäkoski, 2004; Janssen et al., 2005; Streftaris et al., 2005; Cohen, 2006; Fofonoff et al., 2009). Some propose that dispersal could occur via floating plant material drift (Roos, 1979; Koetsier and Bryan, 1989), commercial oysters (Carlton, 1979) and perhaps via birds (Davis, 1957; Green and Figuerola, 2005; Muskó et al., 2008). Reference also has been made to accidental introduction in the Great Lakes by dumping aquaria or via aquatic plants (Mills et al., 1993).

Impact Summary

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Economic/livelihood Negative
Environment (generally) Positive and negative

Economic Impact

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Cordylophora acts as a biofouler by colonizing power station cooling systems and fouling industrial water pipes in Europe and North America (Markowski, 1959; Jenner and Janssen-Mommen, 1993; Jenner et al., 1998; Folino-Rorem and Indelicato, 2005; Escot et al., 2007; Venkastesan and Murthy, 2008; Leppäkoski et al., 2009). Often portions of the colonies can break free at the end of the peak growing season and clog filters in the intake tunnels. Increased salts (chlorides) due to evaporation in bodies of water used for cooling at power plants have created more favourable aquatic habitats for Cordylophora in aquatic ecosystems. Furthermore this hydroid is problematic for shipping, boating and fish farming (Leppäkoski et al., 2009).

Environmental Impact

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Impacts on Habitats

The potential impact on habitats by Cordylophora is extensive since the organism inhabits freshwater and brackish aquatic habitats of various types (Zaiko et al., 2007). Cordylophora can become very abundant and modify habitats by growing on submerged substrata on soft bottoms changing the community structure of soft bottoms (Olenin and Leppäkoski, 1999). Colonies are capable of creating refuges for and from predators and currents and also assist in the accumulation of particulate organic matter (Leppäkoski, 2004). The filamentous structure of Cordylophora colonies may also serve to enhance the settlement and recruitment of dreissenid mussel larvae (Folino-Rorem and Stoeckel, 2006) and the establishment of macroinvertebrates in zebra mussel colonies by providing more surface area (Moreteau and Khalanski, 1994; Folino-Rorem et al., 2006). In freshwater systems, zebra mussel recruitment is highest in areas of increased vegetation (Stan¢czykowska and Lewandowski, 1993), though this depends on plant architecture (Cheruvelil et al., 2002; Kraufvelin and Salovius, 2004). In addition macrophytes also provide macroinvertebrates with refugia from fish predation (Dykman and Hann, 1996; Warfe and Barmuta, 2004, Harrison et al., 2005). Epiphytes, such as the filamentous alga Cladophora or filamentous epifauna such as the hydroid Cordylophora, may enhance macroinvertebrate colonization of mussel colonies. Folino-Rorem et al. (2006) attributed increased zebra mussel settlement on artificial filamentous substrata (hydroid mimics) to an increase in total surface area rather than a preference for filamentous substrata suggesting that settlement of zebra mussel larvae and other macroinvertebrates on aquatic macrophytes may simply be a function of the increase in substrate surface area afforded by these filamentous organisms. These same filaments facilitated the colonization of chironomids and caddisflies; chironomid and caddisfly densities were significantly greater than on control or plates with no filaments (chironomids: p <0.003; caddisflies: p <0.008) (J Stoeckel & N Folino-Rorem, Wheaton College, Illinois, USA, personal communication, 2009).

Cordylophora caspia has been used an indicator species in its ability to absorb metals such as Cd, Zn and Cu at various salinities (Calmano et al., 1992). Future bioassessment protocols should perhaps consider using this hydroid to detect the presence of toxins in freshwater and brackish habitats.

Impacts on Biodiversity 

The ecological impact of Cordylophora needs further exploration though as a sessile raptorial suspension feeder, this predatory hydroid likely modifies aquatic trophic structures by feeding on larval fish prey (Olenin and Leppäkoski, 1999; Folino-Rorem et al., 2007). Cordylophora feeds on chironomids, an important fish food (Menzie, 1981).

The presence of Cordylophora on zebra mussels combined with high macroinvertebrate densities associated with mussel colonies suggests that the impact of Cordylophora on fish prey is likely to be substantial. Understanding how this hydroid may influence densities of benthic macroinvertebrates in dressenid mussel colonies, either positively or negatively, is important for determining the impact of this Ponto-Caspian invader in aquatic benthic communities and thus its potential impact on fish food resources.

In addition to predation, Cordylophora competes with other fouling organisms such as mussels and hydroids (Smit et al., 1993; Jewett, 2005). Cordylophora competes with the bryozoan, Victorella pavida and ciliates for substrate space (Jewett, 2005).

Risk and Impact Factors

Top of page Invasiveness
  • Invasive in its native range
  • Proved invasive outside its native range
  • Has a broad native range
  • Abundant in its native range
  • Highly adaptable to different environments
  • Capable of securing and ingesting a wide range of food
  • Fast growing
  • Has high reproductive potential
  • Gregarious
  • Reproduces asexually
  • Has high genetic variability
Impact outcomes
  • Ecosystem change/ habitat alteration
  • Negatively impacts aquaculture/fisheries
Impact mechanisms
  • Fouling
  • Interaction with other invasive species
  • Predation
  • Rapid growth
Likelihood of entry/control
  • Highly likely to be transported internationally accidentally
  • Difficult to identify/detect as a commodity contaminant
  • Difficult to identify/detect in the field
  • Difficult/costly to control

Similarities to Other Species/Conditions

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Cordylophora colonies occur with other hydroids that are able to tolerate a wide range and/or fluctuations in salinity. In brackish habitats Cordylophora is found with hydroids such as Garveia franciscana (Vervoort, 1964; Arndt, 1984; Ruiz et al., 1999), Gonothyraea loveni and Clavamulticornis (Arndt, 1984).In addition, other co-occurring invasive hydroids found in similar brackish habitats are Maeotias marginata, Blackfordia virginica and Moerisia sp. (Mills and Rees, 2000; Ruiz et al., 2000).

Prevention and Control

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Due to the variable regulations around (de)registration of pesticides, your national list of registered pesticides or relevant authority should be consulted to determine which products are legally allowed for use in your country when considering chemical control. Pesticides should always be used in a lawful manner, consistent with the product's label.

Most attempts to curtail or control the growth of Cordylophora to dateare related to biofouling problems in power plants (Massard and Geimer, 1987; Jenner et al., 1998; Folino-Rorem and Indelicato, 2005; Escot et al., 2007; Venkastesan and Murthy, 2008; Leppäkoski et al., 2009). Methods of eradicating hydroid growth include chlorine and thermal treatments (Rajagopal et al., 2002; Folino-Rorem and Indelicato, 2005). Chlorine and heat combined are advised to curtail growth of Cordylophora (Rajagopal et al., 2002; Folino-Rorem and Indelicato, 2005). Escot et al. (2007) addressed the fouling problems of Cordylophra in a cooling network of the Cartuja'93 technological park in Seville, Spain. They initially attempted to eliminate colonies by manually cleaning pipes. To prevent future colonization, they used a biocide and anti-fouling paint.

Gaps in Knowledge/Research Needs

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There are several of areas of necessary research for the invasive hydroid, Cordylophora:

1. One area is clearly needed in the area of taxonomy. It is critical to know how many species are in the genus Cordylophora as we assess the establishment and associated physiological adaptations pertinent for recruitment and establishment.

2. Identifying the most important factors that enhance the survivorship of recruitment in different habitats (especially those that vary in salinity and food availability) is important is assessing the evolutionary potential of Cordylophora species to adapt to new aquatic ecosystems. This would aid in our understanding of the morphological and ecophysiological responses of Cordylophora species to varying environmental regimes.

3. The ecological impact of this hydroid as a predator and as a provider of filamentous substrate in different aquatic ecosystems would add insight about how this invasive hydroid may potentially alter the biological diversity and community dynamics of aquatic habitats.


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14/08/09 Original text by:

Nadine Folino-Rorem, Wheaton College, Biology Department,, 520 Kenilworth Avenue, Wheaton, Illinois 60187, USA

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